Due to the compact nature of biofilm structures, the presumed reduced physiological state of biofilm bacteria and the protection conferred by biofilm matrix polymers, natural and artificial chemical agents are unable to adequately attack and destroy infectious biofilm populations (Costerton et al., “Bacterial Biofilms in Nature and Disease,” Annu. Rev. Microbiol. 41:435-464 (1987); Hoiby et al., “The Immune Response to Bacterial Biofilms,” In Microbial Biofilms, Lappin-Scott et al., eds., Cambridge: Cambridge University Press (1995)). Increased antibiotic resistance is a general trait associated with biofilm bacteria. When attached, bacteria exhibit a profound resistance, rendering biofilm cells 10-1000 fold less susceptible to various antimicrobial agents than the same bacterium grown in planktonic (free floating) culture. For instance, chlorine (as sodium hypochlorite) an oxidizing biocide considered to be one of the most effective antibacterial agents, has been shown to require a 600 fold increase in concentration to kill biofilm cells of Staphylococcus aureus when compared to planktonic cells of the same species (Luppens et al., “Development of a Standard Test to Assess the Resistance of Staphylococcus aureus Biofilm Cells to Disinfectants,” Appl Environ Microbiol. 68:4194-200 (2002)). Several hypotheses have been advanced to account for the extraordinary resistance of biofilm bacteria to antibiotics including: (i) reduced metabolic and divisional rates exhibited by biofilm bacteria (particularly those deep within the biofilm); (ii) the biofilm EPS matrix may act as an adsorbent or reactant, reducing the amount of agent available to interact with biofilm cells. Additionally, the biofilm structure may physically reduce the penetration of antimicrobial agents by walling off access to regions of the biofilm; (iii) biofilm cells are physiologically distinct from planktonic bacteria, expressing specific protective factors such as multidrug efflux pumps and stress response regulons (Brown et al., “Resistance of Bacterial Biofilms to Antibiotics: A Growth-Rate Related Effect?” J. Antimicrob. Chemotherapy 22:777-783 (1988); Anwar et al., “Establishment of Aging Biofilms: Possible Mechanism of Bacterial Resistance to Antimicrobial Therapy,” Antimicrob. Agents Chemother. 36:1347-1351 (1992); Mah et al., “Mechanisms of Biofilm Resistance to Antimicrobial Agents,” Trends Microbiol. 9:34-39 (2001); Sauer et al., “Pseudomonas aeruginosa Displays Multiple Phenotypes During Development as a Biofilm,” J. Bacteriol. 184:1140-1154 (2002); Stewart, P. S., “Mechanisms of Antibiotic Resistance in Bacterial Biofilms,” Int. J. Med. Microbiol. 292:107-113 (2002); Donlan et al., “Biofilms: Survival Mechanisms of Clinically Relevant Microorganisms,” Clinical Microbiol. Reviews 15:167-193 (2002); Gilbert et al., “The Physiology and Collective Recalcitrance of Microbial Biofilm Communities,” Adv. Microb. Physiol. 46:202-256 (2002); Gilbert et al., “Biofilms In vitro and In vivo: Do Singular Mechanisms Imply Cross-Resistance?” J. Appl. Microbiol. Suppl. 98S-110S (2002)). As detailed molecular studies emerge, it is becoming apparent that each of these factors plays a role in the unusual resistance of biofilms to antimicrobials. Initial treatment is usually effective in killing bacteria only at the margins of biofilm microcolonies. Bacteria deep within these microcolonies are unaffected by the antibiotic and form a nidus for continued dissemination of the infection.
Microbial biofilms in infections and in industrial systems present significant problems due to their recalcitrance to effective treatment.
Detachment is a generalized term used to describe the removal of cells (either individually or in groups) from a biofilm or substratum. Bryers, J. D., “Modeling Biofilm Accumulation,” In: Physiology Models in Microbiology. Bazin et al., eds., Boca Raton, Fla., Vol. 2, pp. 109-144 (1988) categorized four distinct detachment mechanisms by which bacteria detach from a biofilm. These are: abrasion, grazing, erosion, and sloughing. These mechanisms have been described principally from the point of view of the chemical and physical environment acting upon biofilm bacteria. Active detachment as a physiologically regulated event has been hinted at by many authors, but few studies have been performed to demonstrate a biological basis for detachment of bacteria from a biofilm.
One study on the physiological regulation of detachment was carried out by Peyton et al., “Microbial Biofilms and Biofilm Reactors,” Bioprocess Technol. 20:187-231 (1995) on P. aeruginosa. In their work, it was observed that substrate limitation resulted in a decrease in the detachment rate, presumably a result of reducing the growth rate. Allison et al., “Extracellular Products as Mediators of the Formation and Detachment of Pseudomonas fluorescens Biofilms,” FEMS Microbiol. Lett. 167:179-184 (1998) showed that following extended incubation, P. fluorescens biofilms experienced detachment, coincident with a reduction in EPS. In Clostridium thermocellum, the onset of stationary phase has been correlated with increased detachment from the substratum (Lamed et al., “Contact and Cellulolysis in Clostridium thermocellum via Extensive Surface Organelles,” Experientia 42:72-73 (1986)). It has been postulated that starvation may lead to detachment by an unknown mechanism which allows bacteria to search for habitats richer in nutrients (O'Toole et al., “Biofilm Formation as Microbial Development,” Ann. Rev. Microbiol. 54:49-79 (2000)).
The transition from a flowing system to a batch culture system has been observed by many labs to result in biofilm detachment. One lab has observed reproducible detachment of biofilm cells of P. aeruginosa when flow is arrested in a continuous culture system (Davies, D. G., “Regulation of Matrix Polymer in Biofilm Formation and Dispersion,” In Microbial Extracellular Polymeric Substances, pp. 93-112, Wingender et al., eds., Berlin: Springer (1999)).
The release of degradative enzymes has been proposed by others. One such example is found with the gram positive organism Streptococcus mutans which 30 produces a surface protein releasing enzyme (SPRE), shown to mediate the release of proteins from the cell envelope (Lee et al., “Detachment of Streptococcus mutans Biofilm Cells by an Endogenous Enzymatic Activity,” Infect. Immun. 64:1035-1038 (1996)). Boyd et al., “Role of Alginate Lyase in Cell Detachment of Pseudomonas aeruginosa,” Appl. Environ. Microbiol. 60:2355-2359 (1995) showed that over-expression of alginate lyase causes the degradation of alginate. When a mucoid strain of P. aeruginosa was induced to over-express alginate lyase, cells were more easily removed by gentle rinsing from solid medium.
Cell density dependent regulation may also be responsible for the release of enzymes which can degrade biofilm matrix polymers allowing bacteria to disperse from a biofilm. It has been observed at the Center for Biofilm Engineering at Montana State University, USA (Davies, D. G. and Costerton, J. W.) and in the laboratories of Dr. Lapin-Scott at the University of Exeter, UK, that when certain bacteria (including P. aeruginosa) reach high cell densities in biofilm cell clusters, the bacteria often undergo a detachment event. Mutants of P. aeruginosa which lacked the ability to synthesize the quorum sensing autoinducer 3OC12—HSL, were susceptible to detachment following treatment with mild detergent (Davies et al., “The Involvement of Cell-to-Cell Signals in the Development of a Bacterial Biofilm,” Science 280:295-298 (1998)). Other investigators have demonstrated that homoserine lactones may play a role in detachment. Lynch et al., “Investigation of Quorum Sensing in Aeromonas hydrophila Biofilms Formed on Stainless Steel: In: Biofilms—The Good, the Bad and the Ugly, Wimpenny et al., eds. Bioline, Cardiff. pp. 209-223 (1999) reported an increase in detachment of Aeromonas hydrophila from biofilms and Puckas et al., “A Quorum Sensing system in the Free-Living Photosynthetic Bacterium Rhodobacter sphaeroides,” J. Bacteria 179:7530-7537 (1997) reported that homoserine lactone production was negatively correlated with cell cluster formation in Rhodobacter sphaeroides. 
It has been recognized that P. aeruginosa biofilms do not develop into macroscopic biofilm structures in batch culture flasks (at the glass liquid interface). Yet, when medium is pumped continuously through such a flask, (as in a chemostat) a luxurious biofilm develops completely coating the glass surface. When flow is halted in such a system, the biofilm sloughs after a number of days, generally around 72 hrs (Davies et al., “The Involvement of Cell-to-Cell Signals in the Development of a Bacterial Biofilm,” Science 280:295-298 (1998)). The inability of biofilms to develop in batch culture has been observed for a number of gram negative and gram positive bacteria as well as mixed cultures of bacteria. This phenomenon demonstrates that there is some aspect of batch growth that is inhibitory to biofilm development.
During the last stage of biofilm development, the protein profile of bacteria matches more closely the protein profile of planktonic cells than it does biofilm bacteria from the previous stage, denoted maturation II (see FIG. 3 of the current application, and Sauer et al., “Pseudomonas aeruginosa Displays Multiple Phenotypes During Development as a Biofilm,” J. Bacteriol. 184:1140-1154 (2002)).
Due to the compact nature of biofilm structures, the presumed reduced physiological state of biofilm bacteria and the protection conferred by biofilm matrix polymers, current natural and artificial chemical agents are unable to adequately attack and destroy infectious biofilm populations (Costerton et al., “Bacterial Biofilms in Nature and Disease,” Annu. Rev. Microbiol. 41:435-464 (1987); Hoiby et al., “The Immune Response to Bacterial Biofilms,” In Microbial Biofilms, Lappin-Scott et al., eds., Cambridge: Cambridge University Press (1995)).
The present invention is directed to overcoming these and other deficiencies in the art.